1. Labeling Proteins With ICAT Reagent
Typically, protein solutions need to be cleaned up (to remove salts and detergents) by trichloroacetic acid (TCA) precipitation or other compatible cleanup methods. After the salts and detergents have been removed, the sample is dissolved in a minimal volume of labeling buffer. The volume of labeling buffer and the chemical ratio of ICAT reagent can be scaled to achieve an ICAT concentration of approximately 3 mM depending on the amount of sample.
The following protocol is based on labeling 500 μg of protein with each of the two ICAT reagents. If an amount other than 500 μg is used, then the volume of labeling buffer and the chemical ratio of the ICAT reagents can be adjusted proportionally based on the amounts used here.
1) Dissolve the dried or precipitated protein from control and test samples in a minimal volume of freshly prepared labeling buffer.
2) Measure the amount of protein in the control and test samples separately. Save an aliquot (approx 1 μg) from each sample to monitor the labeling efficiency by SDS-PAGE at the end of labeling.
3) Add Tris(2-carboxyethyl) phosphine (TCEP) to a 5-mM final concentration in the protein sample solutions.
4) Add fivefold molar excess of ICAT reagent (see Note 3) to achieve a final concentration of 2.5–3.0 mM in the protein sample solution. Mix well and gently shake the suspension using a tube rocker for 2 h at 37°C in dark.
5) Add fivefold molar excess of dithiothreitol (DTT) to quench the reactions and incubate for 5 min at room temperature. Save an aliquot (approx 1 μg) of proteins from each sample to monitor labeling efficiency.
6) Combine the control and the test labeled samples, and dilute combined sample with 50 mM Tris-HCl (pH 8.3) buffer until the urea is at final concentration of 1 M.
7) Add trypsin 1:100 w/w (trypsin/protein) and incubate the mixture overnight at 37°C.
8) Analyze ICAT labeling efficiency and tryptic digestion by SDS-PAGE followed by silver staining with the saved aliquots from steps 2 and 5.
2 Sample Clean-Up and Fractionation by Strong Cation-Exchange Chromatography
Purification of combined samples using strong cation exchange chromatography depends on sample volume and complexity. For less complex samples or with limited protein available (e.g., ~100 μg of total protein), consideration should be given to preparing a single fraction for avermectin affinity purification using the cation exchange cartridge in the ICAT kit (Applied Biosystems). However, for highly complex samples (i.e., 0.5-1 mg of total protein), fractionation into 25-50 fractions by strong cation exchange (SCX) is recommended.
This section describes the sample fractionation by the SCX chromatography system.
1) Acidify the sample with diluted phosphoric acid to pH 3.0 and load onto the SCX column.
2) Elute ICAT-labeled peptides off of the SCX column using a linear gradient of KCl from 0 to 100% B in 60 min and collect fractions into a glass vial (placed in a 2-mL Micro Tube) at a suitable interval.
3 Affinity Purification of Peptides With Avidin Affinity Cartridge
1) Neutralize each collected cation exchange fraction to pH 7.2 with 500 μL loading buffer.
2) Slowly load (approx 1 drop/s) the neutralized fraction onto the avidin cartridge and collect and save the flow-through into a glass vial.
3) Add an additional 500 μL loading buffer and collect the flow-through into the same tube.
4) Inject 1 mL of wash 1 buffer. Discard the output.
5) Inject 1 mL of wash 2 buffer. Discard the output.
6) Elute the labeled peptides (approx 1 drop/second) with 800 μL of the elution buffer and collect into a glass vial.
7) Repeat steps 1–6 for the rest of the cation-exchange fractions.
4 Cleavage of the Biotin Affinity Tag From the ICAT-Labeled Peptides
1) Dry each fraction completely with a SpeedVac.
2) Prepare the cleaving reagent in a glass vial by combining 95 μL of cleaving reagent A and 5 μL cleaving reagent B.
3) Vortex to mix, and transfer the cleaving reagent to the dried fraction in the glass vials.
4) Cap each glass vial and incubate for 2 h at 37°C.
5) Dry the samples completely with the SpeedVac.
6) Add the appropriate solvent to the dried pellet for MS analysis.
7) Vortex, centrifuge, carefully transfer to the appropriate tube, and store at –80°C until ready for microLC-MS/MS.
5 Isolation and Analysis of Glycopeptides
1) Glycoprotein oxidation: oxidation with sodium periodate converts the cis-diol groups of carbohydrates to aldehydes.
2) Coupling: the aldehydes react with hydrazide groups immobilized on a solid support to form covalent hydrazone bonds. Nonglycosylated proteins are removed.
3) Proteolysis: the immobilized glycoproteins are proteolyzed with trypsin on the solid support. The nonglycosylated peptides are removed by washing, and the glycosylated peptides remain covalently bound to the solid support.
4) Isotope labeling: the α-amino groups of the immobilized glycopeptides are labeled with isotopically light (d0) or heavy (d4) forms of succinic anhydride after the ε-amino groups of lysine are converted to homoarginines.
5) Release: formerly N-linked glycopeptides are released from the solid-phase by PNGase F treatment.
6) Analysis: the isolated peptides are identified and quantified using microcapillary highperformance liquid chromatography electrospray ionization tandem mass spectrometry (microLC-ESI-MS/MS) or microLC separation followed by matrix-assisted laser desorption/ ionization (MALDI) MS/MS.
5.1 Isolation of Glycopeptides
1) Suspend 1 mg of protein in 100 μL of coupling buffer containing 100 mM NaOAc, 150 mM NaCl (pH 5.5).
2) Add sodium periodate solution to a 15-mM final concentration in the sample solutions. Mix well and gently shake the suspension using a tube rocker for 60 min at room temperature in the dark.
3) Remove excess sodium periodate from the sample using a desalting column.
4) Add hydrazide resin equilibrated in coupling buffer to the sample (1 mL gel/5 mg protein). Mix well and gently shake the suspension using a tube rocker for 10–24 h at room temperature in dark.
5) Spin down the resin at 1000g for 10 min, and wash the resin to remove nonglycoproteins three times with 1 mL of 8 M urea/0.4 M NH4HCO3.
6) Suspend the proteins on the resin in 2 M urea/0.1 M NH4HCO3, add trypsin 1:100 w/w (trypsin/protein), and incubate the mixture overnight at 37°C.
7) The peptides can be reduced by adding 8 mM TCEP (Pierce, Rockford IL) at room temperature for 30 min, and alkylated by adding 10 mM iodoacetamide at room temperature for 30 min after reduction.
8) Remove unbound trypsinized peptides.
9) Wash the peptides on the resin with 1 mL of 1.5 M NaCl three times, 80% acetonitrile three times, 100% methanol three times, and 0.1 M NH4HCO3 six times.
10) Suspend resin with 0.5 mL of 0.1 M NH4HCO3, and release N-linked glycopeptides from the resin by incubating with 0.5 μL of peptide-N-glycosidase F at 37°C overnight.
11) Collect the solution from the beads to a clean glass tube, wash the resin twice with 200 μL of 80% CH3CN/0.1% TFA, and combine the supernatant in the glass tube.
12) Dry the combined supernatant completely with a SpeedVac and suspend in 0.4% acetic acid for LC-MS/MS analysis.
5.2 Isotope-Labeling Glycopeptides
1) For isotopic labeling of glycopeptides with succinic anhydride, wash the glycopeptides attached to the resin twice with 15% NH4OH in water (pH > 11.0).
2) Add methylisourea at 1 M in 15% NH4OH (NH4OH/H2O = 15/85 v/v) in 100-fold molar excess over amine groups and incubated at 55°C for 10 min.
3) Wash the beads twice with water, twice with dimethylformamide (DMF)/pyridine/H2O = 50/10/40 (v/v/v), and resuspend in DMF/pyridine/H2O = 50/10/40 (v/v/v).
4) Add succinic anhydride solution to a final concentration of 2 mg/mL. Incubate the mixture at room temperature for 1 h.
5) Wash the mixture three times with DMF, three times with water, and six times with 0.1 M NH4HCO3.
6) Release peptides from the beads using peptide-N-glycosidase F as described.
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Reference
- Walker, J. M. (Ed.). (2005). The proteomics protocols handbook. Humana press.